10 Metabolic shifts in cell proliferation and differentiation

: Alteration of cellular energy metabolism is a principal feature of tumor and stem cells. Here we analyze the metabolic interactions between cancer cells and ﬁ-broblasts in a co-culture model and the metabolic heterogeneity of tumors and metabolic changes in mesenchymal stem cells during adipogenic differentiation based on the ﬂuorescence of the metabolic cofactors NADH, NADPH, and FAD. We registered a metabolic switch from oxidative phosphorylation to glycolysis with slight acidiﬁ-cation of the cytosol in cancer cells in a co-culture model. In the tumor tissue we detected metabolic heterogeneity with more glycolytic metabolism of cancer cells in the stroma-rich zones. The shift of cellular energy metabolism from glycolysis to ox-idativephosphorylationandtheactivationoflipogenesiswereobservedinadipocytes. Data about metabolicalterations incancer andstem cells are important for monitoring the progression of cancers, the development of anticancer drugs and stem cell therapy.


Introduction
Cell metabolism is defined as the sum of the chemical reactions taking place within each cell of a living organism and providing energy for vital processes.It is known that the main way to generate adenosine triphosphate (ATP) for providing living cells with energy is by oxidative phosphorylation (OXPHOS).The metabolic cofactors oxidized flavin adenine dinucleotide (FAD) and reduced nicotinamide adenine dinucleotide (NAD(P)H) are the primary electron acceptor and donor, respectively, in this process.Since these cofactors have a capability for fluorescence, and the fluorescence lifetimes differ for different states, the free or protein-bound, fluorescence intensity and lifetime measurements can be used to monitor the metabolic activity of the cells [1][2][3][4][5].Many enzymes bind to NAD(P)H and FAD in the different metabolic pathways [6].However, in cell proliferation and differentiation an active role in metabolism belongs to glycolysis.Rapidly proliferating cancer cells as well as stem cells tend toward glycolysis because it provides intermediates for the biosynthesis of macromolecules and a fast production of ATP to support their high growth rate [7][8][9].Partial breakdown of glucose through glycolysis and the pentose phosphate pathway provides a compromise between the catabolic generation of ATP and reducing cofactors and production of biosynthetic substrates to meet the cells' anabolic requirements [10].
Multiphoton fluorescence microscopy and fluorescence lifetime imaging (FLIM) are powerful techniques for the non-invasive characterization and long-term monitoring of the functional changes that underlie cellular metabolism.The possibilities for investigating cell metabolism in normal and pathological conditions in vitro and in vivo using these methods have been demonstrated widely [11][12][13][14].
Here we analyze the metabolic interaction between cancer cells and fibroblasts in a co-culture model, the metabolic heterogeneity of tumors in vivo and metabolic changes in mesenchymal stem cells (MSCs) during adipogenic differentiation, based on the fluorescence of the metabolic cofactors NADH, NADPH, and FAD.Cellular metabolism was examined by monitoring the optical redox ratio (FAD/NAD(P)H), the fluorescence lifetime contributions of the free and bound forms of NADH and the bound form of NADPH.Two-photon fluorescence microscopy combined with FLIM was used to analyze this fluorescence in living cells.

Cancer cells and tumor model
HeLa Kyoto, human cervical carcinoma, cells and human skin fibroblasts (huFb) were used in the study.For intracellular pH (pHi) measurements a HeLa cell line stably expressing the cytoplasmic pHi-sensor SypHer2 (HeLa-SypHer2) was used.The huFb were obtained from the Koltzov Institute of Developmental Biology Russian Academy of Science (Moscow, Russia).Genetically transfected cell line was generated and characterized in the Institute of Bioorganic Chemistry RAS (Moscow, Russia).
The cells were cultured in DMEM containing 100 mg/ml penicillin, 100 mg/ml streptomycin sulfate and 10 % fetal bovine serum (FBS) at 37 °C in a humidified atmosphere with 5 % CO 2 .
The protocol for the co-culturing of cancer cells and fibroblasts was modified from that of [15].For co-culturing, the huFb were plated in glass-bottom FluoroDishes in complete DMEM media without phenol red (Life Technologies) and then HeLa Kyoto cells were seeded within 4 hours.The total number of cells per dish in the co-culture was 1 × 10 5 with a 1 : 5 fibroblast-to-cancer cell ratio.In parallel, monotypic cultures were plated at the same quantity as in the corresponding co-culture.The day of plating was defined as a day 0. The day after plating (day 1), the medium was changed to DMEM with 5 % FBS, and afterwards the medium was changed every other day.The cells were analyzed over a period of 5 days.
All animal protocols were approved by the Ethics Committee of Nizhny Novgorod State Medical Academy.Female athymic nude mice of 20-22 g body weight were inoculated subcutaneously in the left flank with HeLa cells (2 × 10 6 in 200 μl PBS).Imaging started 21 days after the cell injection, when the tumors had reached ≈ 10 mm in diameter.Before fluorescence imaging the mice were anesthetized intramuscularly with a mixture of Zoletil (40 mg/kg, Virbac SA, Carros, France) and 2 % Rometar (10 mg/kg, Spofa, Czech Republic), a skin flap over the tumor was surgically opened, and the objective was placed directly on the tumor surface.After image acquisition, the animals were sacrificed by cervical dislocation and the tumors were excised for histopathology.

Stem cells and adipogenic differentiation
All procedures were conducted according to N. K. Koltzov Institute of Developmental Biology Ethic Committee approval.Human bone marrow mesenchymal stem cells (MSCs) were isolated from bone marrow of healthy donors with informed consent according to the Institutional Guidelines under the approved protocol.The MSCs were isolated and characterized as described previously [16].
Differentiation was induced by incubating the MSCs in MesenCult™ Adipogenic Differentiation Medium (Human) (STEMCELL TECHNOLOGIES, Canada).The medium was replaced every 3-4 days over the experimental period of up to 4 weeks.
For microscopic imaging, 4 × 10 5 cells were transferred into a sterile dish with a cover-glass bottom (0.17 mm thick) and incubated for one day until they attached to the glass surface.The cells were imaged before the induction of differentiation (day 0) and on days 5, 12, 19 and 26 subsequently.The cells were washed twice using phosphate-buffered saline, and then placed in FluoroBrite™ DMEM (Gibco) with 10 % FBS and 0.58 mg/ml L-glutamine (PanEco) and 40 U/ml gentamicin.

Two-photon fluorescence microscopy and FLIM
The two-photon exited fluorescence intensity and FLIM images of NAD(P)H and FAD of the cultured cells were obtained on an LSM 710 (Carl Zeiss, Germany) inverted laser scanning confocal microscope.For two-photon fluorescence microscopy and FLIM of NAD(P)H in tumors in vivo a multiphoton tomograph MPTflex (JenLab, Germany) was used.Both systems are equipped with time-correlated single-photon counting (TCSPC) modules (Becker & Hickl GmbH, Germany).
The images of cells in vitro were acquired through a 40×, 1.2 NA water immersion objective.During image acquisition, the cells were maintained at 37 °C and 5 % CO 2 .NAD(P)H and FAD fluorescence was excited with a Chameleon Vision II (Coherent, USA) Ti:Sa femtosecond laser, using an 80 MHz repetition rate and a pulse duration of 140 fs at wavelengths of 750 nm and 900 nm, respectively.Emission was detected in the ranges 455-500 nm for NAD(P)H, and 500-550 nm for FAD.
The images in vivo were acquired through a 40×, 1.3 NA oil immersion objective.NAD(P)H fluorescence was excited at the wavelength of 740 nm using a tunable 80 MHz, 200 fs Ti:Sa laser MaiTai (Spectra Physics, USA) and detected in the range 410-660 nm.Second-harmonic generation (SHG) was exited at 750 nm and detected in the range 373-387 nm to identify collagen in the tumor tissue.
The optical redox ratio FAD/NAD(P)H was calculated from corresponding twophoton fluorescence images of FAD and NAD(P)H after subtracting the background on a pixel-by-pixel basis using ImageJ 1.39p software (NIH, USA).
The analysis of the FLIM data was performed using SPCImage software (Becker & Hickl GmbH, Germany).The lifetime decay curve was fitted to a double-or tripleexponential decay model in the case of NAD(P)H, and to a double-exponential decay model in the case of FAD.The goodness of the fit, the χ 2 value, was 1.2-0.1.An average of 5000-8000 photons were assessed per decay curve for the region of interest (cell cytoplasm).

Metabolic shifts in cancer 10.3.1 Metabolic interaction of cancer cells and fibroblasts
An important role in modifying cancer cell metabolism belongs to the tumor stroma, and, especially, to cancer-associated fibroblasts (CAFs).Multiple studies report on the CAF's ability to promote tumorigenesis and to regulate cancer cell motility and stemness through the secretion of a number of growth factors, hormones and cytokines [17].Metabolic coupling between cancer cells and CAFs occupies a special role in these processes because metabolic reorganization underlies further changes leading to adaptation of the cancer cells.Interestingly, in some tumors CAFs undergo metabolic alterations and switch their metabolism to aerobic glycolysis when they interact with cancer cells, while the cancer cells themselves remain oxidative, the phenomenon being known as the "reverse Warburg effect" [18].
To estimate the overall cellular metabolic activity, measurements of the fluorescence intensity of FAD to NAD(P)H were performed in the co-culture of cancer cells and fibroblasts and in the corresponding monocultures.Cancer cells and fibroblasts in the co-culture were identified by their cellular morphology.Cancer cells are much smaller than fibroblasts, have a round shape and large nucleus with a thick cytoplasmic layer.The fibroblasts have a typically spindle-shaped morphology, much greater size and a larger volume of cytoplasm compared to the cancer cells.
Monitoring of the redox ratio FAD/NAD(P)H in the co-culture showed that in the cancer cells the redox ratio significantly decreased on day 5 of co-culturing, indicating an increase in metabolic activity (Fig. 10.1 (a)).Meanwhile, the fibroblasts showed a gradual increase of their redox ratio from day 1 to day 5, indicating a probable metabolic shift toward OXPHOS [19].In the monocultures the redox ratio in the fibroblasts was higher than in the cancer cells, which testifies to higher oxidative metabolism in the stromal cells, but did not change with time in either cell type (1.4 vs. 0.35, p = 0.000000) (Fig. 10.1 (b)) [19].
Our results are consistent with the data by Ostrander et al., who showed that the MCF-10A and HMEC cell lines of normal mammary epithelial cells had an increased redox ratio when compared to nine different breast cancer cell lines [20].Martinez-Outschoorn et al. also detected increased mitochondrial activity in fibroblasts when compared with the MCF-7 cell line in monoculture [15].Similar results were reported by de Andrade Natal et al. for biopsies of breast cancer patients [21].Cancer cells, where aerobic glycolysis predominates, display a decreased redox ratio FAD/NAD(P)H, as has been demonstrated for different cancer cell lines and tumors [14,20].
As expected for FLIM, the fluorescence decay curves for NAD(P)H and FAD were best fitted to a double-exponential decay model, indicating the presence of two distinctly different lifetimes for the free and protein-bound forms of the cofactors.The fluorescence lifetimes of the free (τ 1 ) and protein-bound (τ 2 ) NAD(P)H were measured to be ≈ 0.4 and ≈ 2.6 ns, respectively.For the free (τ 2 ) and protein-bound (τ 1 ) FAD fluorescence lifetimes were ≈ 2.7 and ≈ 0.4 ns, respectively.Fluorescence lifetimes documented in the literature lie in the range of 0.2-0.5 ns for free NAD(P)H, 1.8-2.7 ns for protein-bound NAD(P)H, 0.15-0.4ns for protein-bound FAD and 2-2.8 ns for free FAD [5,14,22].Therefore, the values recorded in our study agree well with singular data.
We have found no difference in the fluorescence lifetimes between cancer cells and fibroblasts of one cell type in the process of co-cultivation in vitro.However in work by Skala, increased fluorescence lifetimes of free and protein-bound NAD(P)H and free FAD, and decreased fluorescence lifetimes for protein-bound FAD in normal tissue compared to oral precancerous tissue were detected in vivo [23].McGintly et al. found an increased mean fluorescence lifetime for NAD(P)H in colon cancer but a decreased lifetime for precancerous colon lesions compared to the normal colon [24].
The relative contribution of free NAD(P)H (a 1 ) in cancer cells co-cultured with fibroblasts increased from 76.9 % on day 1 to 79.45 % (p = 0.000001) on day 2, and remained at that increased level during whole period of co-culturing.The relative contribution of free FAD (a 2 ) in these cells increased on day 3 from 27.9 to 32.4 % (p = 0.000000), and then did not change.The observed changes in the relative contributions of free NAD(P)H and FAD testify to an increased bias toward a glycolytic metabolism.By contrast, for the fibroblasts in the co-culture, the relative contributions of free NAD(P)H and FAD gradually decreased starting from day 2, indicating a shift toward oxidative metabolism (Fig. 10.2).All cells in the population displayed the described changes.
In monocultures of cancer cells and fibroblasts the relative contributions of the cofactors were fairly stable throughout the 5 days of cultivation (≈ 76 % for free NAD(P)H and ≈ 30 % for free FAD) without any statistical difference for the NAD(P)H and only a slight difference for FAD (p = 0.000011).
Previously, in vivo, a decreased contribution of protein-bound NAD(P)H and FAD had been demonstrated for precancer development [24].A decreased amount of NAD(P)H and a decreased free/bound NAD(P)H ratio were also shown in oral squamous carcinoma cells when compared with nonmalignant cells [13].A glycolytic phenotype of cervical cancer had been demonstrated for cell cultures and human tumors.For example, Rossignol et al. reported that HeLa cells generate energy predominantly by glycolysis, but can change ATP generation to exclusively OXPHOS when the availability of glucose is limited [25].It was found by Herst et   cells that rely preferentially on OXPHOS in highly oxygenated conditions became more glycolytic upon hypoxia [26].Glycolytic metabolism of HeLa cells was also shown for tumor spheroids [27].At the same time, fibroblasts retain a significant level of OXPHOS even if the glucose level is high [28,29].Nevertheless, there have been very few studies in this field performed on the co-culture model.For example, a higher glucose uptake and increased lactate production in co-cultures of MCF-7 and CAFs compared with monocultures were demonstrated by Brauer et al., but in that work the cancerous and the normal cells were not separated [30].Martinez-Outschoorn et al. demonstrated the reversed Warburg effect (OXPHOS in cancer cells and glycolysis in fibroblasts) in co-cultures of MCF7 cancer cells and fibroblasts [15].Therefore, the FLIM measurements of the relative contributions of protein-bound and free NAD(P)H and FAD in cancer cells and fibroblasts showed a switch of the HeLa cells toward a glycolytic phenotype and a switch of the huFb toward OXPHOS as a result of co-cultivation.
Although there are some papers demonstrating the dependence of the metabolic state on the stage of cell culture growth, in particular a decrease in the fluorescence lifetime of both free and protein-bound NADH and the contribution of protein bound NADH at higher cell densities [31,32], the observed metabolic changes in our study with co-culture were most probably not related with this.Control monocultures were grown under similar conditions and had the same density as co-culture, but did not show any fluctuations of redox ratios and fluorescence lifetimes.
Analysis pHi in cancer cells in a co-culture model was performed using a new ratiometric genetically-encoded pH sensor SypHer2.SypHer2 is an improved pH sensitive ratiometric indicator based on the cpYFP fluorophore.SypHer2 has two excitation peaks, at 420 nm and 500 nm, and one emission peak at 516 nm [33,34].The excitation peak at 420 nm decreases with pH proportionally to the increase in the peak at 500 nm, allowing ratiometric (dual excitation) imaging of intracellular pH in living cells.
In monoculture of cancer cells stably expressing the sensor, the fluorescence ratio I 500 /I 420 did not change with time, indicating a stable pHi (Fig. 10.3).
On day 1 of co-cultivation with fibroblasts, the SypHer2 ratio had already started to show a reduction compared to that in the monoculture (p = 0.028) because the pHi became more acidic.The lower SypHer2 ratio remained throughout further cultivation.A minor acidification of the cytosol of cancer cells co-cultured with fibroblasts corresponds well with metabolic shift to glycolysis shown by NAD(P)H and FAD fluorescence imaging: decreased pHi is associated with increased lactic acid production in conditions of increased glycolysis.The combination of lactic acidosis and decreased cytosolic pH is a specific feature of cancer cells with high rates of glycolysis [35].It is known that the accumulation of lactate during tumor development may affect the pHi and drive metabolic changes [36].At the same time, for successful realization of glycolysis, an elevated pH level is required [37].
It should be noted that there is a great diversity of metabolic phenotypes among tumors, and besides the above mentioned "Warburg type" and "reverse Warburg type" there are tumors where both cancer cells and stroma are glycolytic or nonglycolytic [38].Our HeLa-huFb system demonstrated metabolic behavior similar to Warburg type tumors, where cancer cells use increased glycolytic and stromal cells tend toward increased nonglycolytic metabolism.

Metabolic heterogeneity of tumors
The great degree of metabolic heterogeneity, both inter-and intratumoral, and the high plasticity of cancer cells significantly complicate the development of effective therapeutic strategies and result in nonuniform response to therapy.In general, the metabolism of cancer cells is more flexible compare to normal cells, which gives them the opportunity to survive in unfavorable conditions.Such factors as hypoxia, glucose deprivation, lactate accumulation and extracellular acidosis contribute a great deal to tumor progression through forming a specific microenvironment and thus they determine, to a great extent, the metabolic phenotype [39].It has been demonstrated that some cancer cells can reversibly switch between fermentation and oxidative metabolism, depending on the absence or the presence of glucose and oxygen and other environmental conditions [40].Therefore, cancer metabolism is neither as homogenous nor as reproducible as initially suspected.Rather, the metabolic activity of cancer cells is a complex, heterogeneous, and nuanced process that may be a key to successful treatment.
Fluorescence lifetimes of free and protein-bound forms of NAD(P)H and their contributions were measured in vivo in the HeLa tumors in nude mice [41].Fluorescence intensity of FAD was very low in the tumor tissue, which did not allow fluorescence lifetime measuring of this cofactor.
Macroscopically, the tumors had a multinodular, fleshy appearance with blood vessels, plus yellowish and red-colored areas.
The fluorescence lifetimes of the free (τ 1 ) and protein-bound (τ 2 ) NAD(P)H measured in vivo in cancer cells were 0.47 ± 0.8 ns and 2.3 ± 0.3 ns, respectively.
The relative contributions of free (a 1 ) and protein-bound (a 2 ) NAD(P)H in cancer cells were different in different tumor sites (Fig. 10.4).In some areas the relative contribution of free NAD(P)H was 75.5 ± 2.4 %, while in other zones it increased to 80.8 ± 2.7 % (p = 0.000001), indicating heterogeneity of cellular metabolism within a tumor node.Intravital fluorescence intensity and SHG microscopy and subsequent histological examination showed that the areas with greater contribution of free NAD(P)H, and consequently more glycolytic metabolism, were enriched with connective tissue fibers and included not numerous cancer cells.Whereas more oxidative areas, with greater contribution of the protein-bound NAD(P)H, had more dense cellular structure with low content of fibrotic stroma.Therefore, we demonstrated for the first time the metabolic heterogeneity in the HeLa tumor in vivo using FLIM.Cancer cells in cell-rich and stroma-rich areas differed in their energy metabolism.In the literature, metabolic heterogeneity has been shown for various tumor types [42].It is considered that all cancerous cells that make up a tumor do not behave in a uniform fashion since genes can be regulated at the single cell level [43].
It is known that cancer-associated extracellular matrix (ECM) actively contributes to histopathology and behavior of tumors.Meanwhile, the interplay between ECM mechanics and energy metabolism of cancer cells is not fully understood.Emerging evidence consistently indicates that ECM stiffness and increased ECM deposition may influence cellular metabolism, thereby promoting both cell proliferation and survival [44], as well as facilitating oncogenic transformation and tumor metastasis [45].It is supposed that the stiffened ECM, often enriched for type I collagen, promotes glucose uptake and aerobic glycolysis through the mechanical activation of the protumorigenic signaling pathways (PI3K/Akt) [46].

Metabolic shifts in stem cells
Energy metabolism is known to be an important regulator of the ability of stem cells to undergo both differentiation and self-renewal.We investigated the metabolic changes in living MSCs during adipogenic differentiation [47].
MSC differentiation was assessed by noting morphological changes and the development of lipid vacuoles.On days 5 and 12 of differentiation the MSCs had a spindleshaped morphology; the cell population was homogeneous.By days 19 and 26, 14 % and 36 %, respectively, of the total number of cells had become polygonal in morphology and contained lipid vacuoles.
To estimate the general level of metabolic activity of the cells during adipogenic differentiation, the fluorescence intensities of NAD(P)H and FAD were measured and represented as their redox ratio (FAD/NAD(P)H).No significant changes in the redox ratio values were detected until 19 days after the induction of differentiation.On days 19 and 26 a decrease in the redox ratio could be observed (Fig. 10.5).
The changes in the cofactor fluorescence and, consequently, in the redox ratio for differentiated adipocytes may be associated with both energy metabolism and the biosynthesis of lipids.
The optical redox ratio is sensitive to the balance between the rates of ATP consumption and glucose catabolism in a cell.Undifferentiated MSCs produce ATP primarily through glycolysis, and this is accompanied by NADH production, while those undergoing differentiation display a shift towards OXPHOS for their energy needs.It is generally accepted that glycolytic cells show decreased redox ratios, while in oxidative cells the ratio increases [48].
Since a metabolic shift to OXPHOS is expected during the differentiation of stem cells and that would lead to an increase in the redox ratio, we speculated that the observed decrease of the ratio resulted from the accumulation of NADPH in the process of lipogenesis.
To estimate the contribution of energy metabolism and lipogenesis in the metabolic profile, a separate analysis of NADH and NADPH is required.To separate NADH and NADPH in differentiating stem cells we used FLIM with a three-exponential fitting of the fluorescence decay curves, using the method developed be Blacker et al. [49].In 2014, Blacker et al. were the first to use FLIM for the separation of NADH and NADPH fluorescence in live cells (HEK293) and tissues (mammalian cochlea).Using numerical methods in combination with biochemical manipulations on the cells, the intracellular fluorescence lifetimes of bound NADH and bound NADPH were evaluated to be 1.5 ± 0.2 and 4.4 ± 0.2 ns, respectively [49].
Since the differentiation of the MSCs may result in changes in the fluorescence lifetimes and the amplitudes of the free and the two bound forms of NAD(P)H, it would be natural to use the three fluorescence lifetime components in a fitting model of the fluorescence decay.
It was found that the contribution of free NADH did not change up to day 19.A statistically significant decrease in the short fluorescence lifetime contribution was shown in differentiated cells (days 19 and 26) in the regions of cytoplasm at the cell periphery as compared with undifferentiated MSCs (day 0) (Fig. 10.6 (a)).
The reverse trend was observed in the contribution of bound NADH on days 19 and 26.It is interesting that the fluorescence lifetime contribution of bound NADH increased only in the regions of cytoplasm at the periphery of the cells.The statistically significant decrease in the contribution of the bound form of NADH on day 12 of differentiation is probably associated with the redistribution of bound NADH to bound NADPH (Fig. 10.6 (a)).
The contribution of bound NADPH was higher at all time points of adipogenic differentiation when compared with undifferentiated MSCs.The maximum elevation of bound NADPH was detected in differentiated cells (days 19 and 26) without, however, any differences between the zones of the cytoplasm at the cell periphery and around the lipid vacuoles (Fig. 10.6 (a)).
It can be seen from Fig. 10.6 (b) that the distribution of the a boundNADH /a boundNADPH ratio in adipocytes is inhomogeneous, varying from 0 in the areas of the cytoplasm next to lipid vacuoles (which indicates a contribution to the ratio only from NADPH), to 2 in the other parts of the cytoplasm.
Therefore, the FLIM data, processed with the three-exponential fitting model, on the increase in the contribution of protein-bound NADH in the cytoplasm at the cell periphery testify to the metabolic switch from glycolysis to OXPHOS during the differentiation of stem cells.The rise in the contribution of bound NADPH is probably associated with activation of fatty acid biosynthesis, where NADPH is involved.
Similar to our work, Quinn et al. showed a decrease in the optical redox ratio associated with lipogenesis during adipogenic differentiation [50].They suggested that adipogenic differentiation, at least in vitro, is associated with an increase in flux through the metabolic pathways using enzymes expressed in the mitochondria.Citrate can then be shuttled out of the mitochondria and cleaved in the cytosol so that acetyl-CoA can be used as a carbon supply for fatty acid synthesis.This process would require the conversion of NAD + to NADH by the LipDH-containing pyruvate dehydrogenase complex (PDHC).Carbon storage in lipid droplets through de novo fatty acid synthesis would probably not occur when substantial ATP production is necessary, and consequently the reduced form of NADH may accumulate.The remaining cytosolic oxaloacetate can be converted to pyruvate by malic enzyme (NADPH is also produced during its conversion) for use in long chain fatty acid synthesis [51].A decreased redox ratio in metabolically active, adipogenically differentiated cells was also detected in [12].König et al. showed an increase in the bound forms of NAD(P)H during the differentiation of stem cells into adipocytes [52], Stringari et al. demonstrated a higher free/bound NADH ratio in undifferentiated neural progenitor and stem cells than in differentiated neurons, and that stem cells followed a metabolic trajectory from a glycolytic phenotype to an OXPHOS phenotype through the different stages of differentiation [53].
Our results about lower contribution of NADPH compared to NADH to the total fluorescent signal are consistent with other studies.Blacker et al. showed that the concentration of bound NADPH decreased ≈ 3-fold in HEK293 cell lines [49].Quinn et al. demonstrated on pancreatic islet and porcine heart that the intracellular concentration of NADH was 33.7 ± 13.1-fold higher than NADPH [50].Klaidman et al. showed that the concentration of the fluorescent reduced NADH was 5 times greater than that of the fluorescent NADPH in mouse hippocampus [54].Moreover, the enhancement of mitochondrial NADH quantum yield due to environmental effects has been estimated to be a factor of 1.25-2.5 greater than that of NADPH [55].Protein binding can also affect the fluorescence quantum yield of NADH, with up to a 10-fold increase in the fluorescence intensity of mitochondrial protein-bound NADH relative to free NADH [56].
To the best of our knowledge, the changes in the contribution of bound NADPH in MSCs during adipogenic differentiation have never previously been measured using FLIM with three-exponential fitting of the fluorescence decay curves.However, using the contribution of bound NADPH and NADH for the study of phenotypic changes requires further investigation and appropriate metabolic measurements.

Conclusions
Effective control of cell proliferation and differentiation is a great challenge because of the complex relationships between the different signaling pathways, the extracellular microenvironment, and the metabolic requirements of the cell.There are many methods for assessing the metabolic status of cells, including immunocytochemistry or immunohistochemistry for metabolic markers (for example, the monocarboxylate transporters MCT1 and MCT4); measuring glucose uptake; lactate production and oxygen consumption; evaluation of the key glycolytic enzymes (for example, hexokinase-II, phosphofructokinase and pyruvate kinase) and the microarray analysis of glycolytic gene expression.However, these methods are invasive, require the introduction of exogenous labels into the cells and do not provide complete information about the physiological changes in the living cells during their growth, proliferation and differentiation.In contrast, multiphoton fluorescence microscopy combined with FLIM is an emerging modality that permits repeated nondestructive measurements of the components within living cells.To date, multiphoton microscopy of NAD(P)H and FAD provides an established technique for metabolic imaging in vitro and in vivo.The data about metabolic changes in tumor-stroma co-evolution are important for the development of anticancer drugs targeted on metabolic pathways in both cancer and adjacent stromal cells.Understanding the metabolic peculiarities of stem cells and the changes accompanying the differentiation process is extremely important for the development of new treatment strategies in regenerative medicine and stem cell therapy.

Fig. 10 . 3 :
Fig. 10.3: Analysis of pHi in cancer cells in monoculture and in co-culture with fibroblasts using genetically encoded sensor SypHer2.Ratiometric images I 500 /I 420 (a), SypHer2 ratio in monoculture (b) and in co-culture (c) from days 1 to 5 of culturing.* statistically significant difference from monoculture on the same day, p ≤ 0.05.

Fig. 10 . 6 :
Fig. 10.6:The dynamic of fluorescence lifetimes contributions of free NADH, and protein-bound forms of NADH and NADPH in undifferentiated MSCs and cells at all stages of differentiation (a).M ± SD. * statistically significant difference with day 0. p values are shown.The distribution of NADH and NADPH in undifferentiated (day 0) and adipogenically differentiated MSCs (day 26) (b).False color-coded images of the a bound NADH /a bound NADPH ratio.Field of view is 213 µm × 213 µm (512 × 512 pixels).V: vacuoles, N: nucleus, P: periphery of cell.